genetic dissection of an exogenously induced biofilm in ......genetic dissection of an exogenously...

Genetic Dissection of an Exogenously Induced Biofilm inLaboratory and Clinical Isolates of E. coliSasan Amini, Hani Goodarzi, Saeed Tavazoie1*

Department of Molecular Biology and Lewis-Sigler Institute for Integrative Genomics, Princeton University, Princeton, New Jersey, United States of America

Abstract

Microbial biofilms are a dominant feature of many human infections. However, developing effective strategies forcontrolling biofilms requires an understanding of the underlying biology well beyond what currently exists. Using a novelstrategy, we have induced formation of a robust biofilm in Escherichia coli by utilizing an exogenous source of poly-N-acetylglucosamine (PNAG) polymer, a major virulence factor of many pathogens. Through microarray profiling ofcompetitive selections, carried out in both transposon insertion and over-expression libraries, we have revealed the geneticbasis of PNAG-based biofilm formation. Our observations reveal the dominance of electrostatic interactions between PNAGand surface structures such as lipopolysaccharides. We show that regulatory modulation of these surface structures hassignificant impact on biofilm formation behavior of the cell. Furthermore, the majority of clinical isolates which producedPNAG also showed the capacity to respond to the exogenously produced version of the polymer.

Citation: Amini S, Goodarzi H, Tavazoie S (2009) Genetic Dissection of an Exogenously Induced Biofilm in Laboratory and Clinical Isolates of E. coli. PLoSPathog 5(5): e1000432. doi:10.1371/journal.ppat.1000432

Editor: Mark Achtman, University College Cork, Ireland

Received November 14, 2008; Accepted April 15, 2009; Published May 15, 2009

Copyright: � 2009 Amini et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permitsunrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: ST was supported by grants from the NSF (CAREER), DARPA, NHGRI, NIGMS (P50 GM071508), and the NIH Director’s Pioneer Award (1DP10D003787-01). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing Interests: The authors have declared that no competing interests exist.

* E-mail: [email protected]

Introduction

Biofilms are an integral component in the life-cycle of many

microorganisms. Compared to their planktonic complement,

however, bacterial biofilms have remained poorly understood,

mostly due to the inherent complexities associated with biofilm

studies, including spatial heterogeneity of the biofilm structure,

longer generation time, and uncharacterized growth parameters [1].

Bacterial biofilms are characterized by the presence of an

extracellular polymeric matrix, which encases the cells. The

physicochemical properties of this matrix, including its charge,

porosity, and architecture are prominent determinants of biofilm

lifestyle. The matrix, for example, could act as a protective barrier

by interacting with large, charged, or reactive biocidal agents and

neutralizing them [1]. One major component of matrix in various

bacterial species is a homopolymer of N-acetylglucosamine. In

fact, poly-N-acetylglucosamine (PNAG) is the major virulence

factor of Staphylococcus epidermidis [2]. There is increasing evidence

that this polysaccharide is produced by a variety of other

pathogens including Bordetella, Yersinia, Staphylococcus, Actinobacillai,

and certain pathogenic Escherichia coli strains as well. It was

reported that enzymatic hydrolysis of poly-N-acetylglucosamines

disrupts biofilm formation by Yersinia pestis, Pseudomonas fluorescens,

Aggregatibacter actinomycetemcomitans, pathogenic E. coli strains, and

various Bordetella species [3–7]. This suggests that PNAG is a

critical component of the biofilm structure made by all these

bacteria. Furthermore, a recent study showed that most E. coli

strains isolated from urinary tract and neonatal bloodstream

infections possess the pga locus required for PNAG biosynthesis,

and almost all of them produce immunologically detectable levels

of PNAG [8].

Involvement of PNAG-based biofilms in the pathogenesis of

various bacterial species makes it an important phenomenon to study

[4,8–11]. Even though the properties of PNAG-based biofilms have

been extensively studied in Staphylococcus species [2,12], the existence

of a PNAG-based matrix in biofilm structures from other species,

including E. coli, has been reported only recently [3,13], and is not as

well characterized as it is in Staphylococcus species. However, there are

some features of PNAG-based biofilm like spatial distribution of the

cells in PNAG-based biofilms that are better studied in E. coli [14]. In

E. coli K-12, expression of PNAG biosynthesis genes is not high

enough to support the formation of a robust biofilm structure under

laboratory conditions [15], complicating the analysis of this

phenotype. Therefore, in order to study the genetic basis of

PNAG-based biofilm formation, we decided to enhance this

phenotype in E. coli K-12 by either increasing the level of

endogenous PNAG or providing an exogenously produced form of

PNAG.

PNAG production in E. coli can be enhanced by manipulation

of the genetic elements involved in pga locus regulation [16,17].

For example, E. coli csrA mutants overproduce the PNAG polymer

[13,17]. However, csrA is a master regulator of the carbon storage

system, and csrA mutants show highly pleiotropic phenotypes. An

alternative approach for enhancing PNAG-based biofilm forma-

tion would be to use a functionally active exogenous source of

PNAG to induce biofilm formation in E. coli. In the csrA mutant

background, cells secrete PNAG polymer into the growth media

[18]. Therefore, the spent media from the csrA mutant culture can

be used as a potential source of PNAG for inducing biofilm

formation.

We observed that application of exogenously produced PNAG,

isolated from DcsrA cells saturated culture of csrA mutants, led to

PLoS Pathogens | www.plospathogens.org 1 May 2009 | Volume 5 | Issue 5 | e1000432

robust biofilm formation in the E. coli K-12 background. This

provided us a unique opportunity to extensively characterize the

underlying genetics of the PNAG-based biofilm formation

phenomenon. Given our observations, we favor a model in which

electrostatic interactions between this polysaccharide and cell

surface structures, such as lipopolysaccharide (LPS), are critical for

PNAG-induced biofilm formation by E. coli. We also show that

although response to PNAG polymer is a purely structural

phenomenon, it can be modulated by multiple pathways including

LPS biosynthesis, the acid tolerance system, capsule biosynthesis,

and regulation of cell morphology.

Results

Biofilm-Inducing Activity of Secreted PNAG in SpentMedia of DcsrA Cells

Escherichia coli csrA mutants secrete poly N-acetylglucosamine

polysaccharide into the culture medium [18]. In order to see

whether the secreted polysaccharide is functional, we grew DcsrA

cells to stationary phase, discarded the cells, and added the cell-

free spent media to wild-type MG1655 cells in the presence of

fresh media. Interestingly, we observed that the cell-free spent

media of saturated DcsrA cultures made wild-type cells form a

biofilm on a rapid time-scale (Figure 1, compare i and ii). Presence

of a carbon source, as expected, was required for formation of

mature visible microcolonies by living wild-type cells (Figure 1,

compare iv and vii to v and vi). Synthesis of PNAG in E. coli

requires the gene products of the pgaABCD operon [13,17]. To test

whether the observed biofilm-inducing activity is associated with

the secreted PNAG (sPNAG), we generated four double mutants

each harboring csrA deletion together with the deletion of one of

the four genes present in the pga locus. There was no detectable

biofilm-inducing activity in the cultures of any of the four double

mutants. Furthermore, the biofilm-inducing activity was lost after

treatment of DcsrA culture spent media with Dispersin B, an

enzyme that specifically cleaves PNAG [3,5,19], confirming that

the biofilm inducing factor is an N-acetylglucosamine containing

polysaccharide. Differential up-regulation of the pga locus

transcription between the DcsrA and wild-type cells was also

confirmed by a reporter assay (Figure S1).

In order to confirm that the observed biofilm formation

phenotype is specific to sPNAG and rule out the possibility of

involvement of other biofilm-inducing agents that might be

present in the DcsrA spent media, we purified sPNAG from spent

media. As part of the purification steps, sPNAG was treated with

various enzymes, including DNase, RNase, a-amylase, and

Proteinase (see Materials and Methods section). The purified

sample showed identical biofilm-inducing activity, suggesting that

sPNAG is sufficient for inducing biofilm formation. The purified

polysaccharide was also characterized by mass spectrometry. As

shown in Figure 2, almost all prominent molecules found on the

mass spectrum corresponded to N-acetylglucosamine oligomers

with different levels of acetylation or to their monomers. In S.

epidermidis, deacetylation of PNAG polymer introduces positive

charges in the otherwise neutral polymer [20]. Our results also

indicate that a considerable fraction of sPNAG building units is

deacetylated, which should leave a net positive charge on the

polymer. Purified PNAG isolated from Staphylococcus aureus strain

MN8m [21] showed similar biofilm-inducing activity when

applied to wild-type MG1655 cells, further confirming that PNAG

is sufficient for the observed biofilm-inducing activity. The

presence of various identical peaks in the mass spectra of PNAG

from E. coli and S. aureus (Figure S2) indicates that they are closely

related molecular species.

The response of wild-type E. coli cells to sPNAG was so fast that

we decided to study the early stages of the process. Using time-

lapse microscopy (Video S1), we observed that wild-type cells

started seeding microcolony structures on a glass slide in less than

an hour of exposure to sPNAG. The pace of microcolony

formation observed here was much faster than the previously

reported behavior by csrA mutants [15]. The microcolonies

expanded in size due to both growth of pre-existing cells and

continued incorporation of new cells. No similar activity was

observed in the absence of sPNAG (Video S2). These results show

that sPNAG enhances both cell-cell and cell-surface interactions.

SEM images of the biofilm structures formed by wild-type cells in

the presence of sPNAG also confirmed the presence of an

extracellular matrix encasing the cells (Figure S3).

Genome-Wide Identification of Loci Involved in sPNAG-Based Biofilm Formation

We used a microarray-based genetic footprinting strategy [22]

to study the genetic basis of sPNAG-mediated biofilm formation.

Since formation of biofilm by wild-type cells in the presence of

sPNAG is robust and fast, the phenotype is highly amenable to

genetic analysis. Starting from a close to saturation Tn5-based

library of approximately 56105 independent transposon inser-

tional mutants, generated in wild-type E. coli MG1655, we devised

a selection strategy to enrich for mutants defective in responding to

sPNAG. Roughly 1010 cells of the abovementioned library were

exposed to sPNAG in LB medium. After 12 hours of incubation,

cells which were present in the liquid phase of the culture and were

not part of the biofilm were isolated, grown up to log phase, and

transferred to a new container with fresh media and fresh sPNAG,

in order to enrich for mutants impaired in responding to sPNAG.

After four rounds of serial enrichment, no visible biofilm formation

activity was present in the enriched population. A schematic

representation of the enrichment procedure is shown in Figure 3.

To quantify the contribution of different loci to this impaired

biofilm formation phenotype, the insertion sites of the transposon

Author Summary

Both in the wild and in the clinical setting many bacterialspecies live within surface-attached communities calledbiofilms. It is still unclear the extent to which the biofilmlifestyle and its associated phenotypes, such as hyper-tolerance to antimicrobial agents, can be attributed tostructural characteristics of the biofilm community or tointrinsic biofilm-specific physiological programs. In orderto address this longstanding question, we focused onpoly-N-acetylglucosamine (PNAG)–based biofilms, a clini-cally relevant phenotype of many bacterial pathogens,including E. coli. Instead of working in a biofilm-permissivegenetic background, in which the timescale of biofilmformation is slow, we applied the functionally activesecreted version of the PNAG exo-polysaccharide (sPNAG)to wild-type E. coli cells, generating robust biofilms on thetimescale of hours. In this way, we managed to uncoupleupstream regulatory processes and matrix preparatoryphase of biofilm formation, focusing specifically on thelatter part. By using a powerful genome-wide technology,we identified the genes and pathways involved in sPNAG-based biofilm formation. Our results revealed thatstructural interactions between sPNAG and surface struc-tures such as lipopolysaccharides are the crucial determi-nants of biofilm formation and that multiple pathwaysincluding acid-tolerance, capsule biosynthesis, and regu-lation of cell morphology modulate this phenotype.

Genetic Basis of PNAG-Based Biofilm Formation


in the enriched population were mapped using a microarray-based

approach [22]. The histogram in Figure 4A shows the normalized

average output of the hybridization data from two experimental

replicates. The z-score for each ORF is indicative of the

abundance of transposon insertion events in that ORF (or its

vicinity) in the enriched population of mutants. More detailed

Figure 1. Secreted PNAG induces biofilm formation. (i) Wild-type cells in LB+sPNAG. (ii) Wild-type cells in LB. (iii) DrfaY cells in LB+sPNAG. (iv)Wild-type cells in PBS. (v) Wild-type cells in PBS+.2%Glucose+sPNAG. (vi) Wild-type cells in PBS+.2% Lactose+sPNAG. (vii) DlacZ cells inPBS+sPNAG+lactose. In the experiments carried out in the presence of lactose, since there was no pre-induction phase with lactose, the observed celldensity was lower.doi:10.1371/journal.ppat.1000432.g001

Figure 2. ESI LTQ OrbiTrap mass spectrum of digested sPNAG, acquired in positive mode. Purified sPNAG was digested with Dispersin Bin water and analyzed by mass spectrometry after dialysis. For simplicity, only the most prominent peaks are labeled with their m/z and z values(where z is the charge). For each labeled peak, molecular composition was schematically illustrated by red and green circles representing acetylatedand non-acetylated saccharide units, respectively (e.g., two red and three green circles corresponds to a pentamer with two N-acetylglucosamine andthree glucosamine residues). For all identified molecules, the ion-pairs corresponding to the singly-charged protonated ions and their dehydrationproducts were detected (e.g., 383.17 and 365.16). As shown in the figure, all prominent peaks correspond to a partially de-acetylated N-acetylglucosamine oligomer or the monomers. Almost all unlabeled peaks correspond to one member of some dehydrated-nondehydrated ion pairs,doubly charged versions of some of the expected molecules, or some adducts. A complete list of m/z values for all potential mono- andoligosaccharides species that could be generated from an incomplete digestion of a PNAG sample with all possible acetylation patterns is also givenin Table S1, as a reference.doi:10.1371/journal.ppat.1000432.g002



information regarding the calculation of z-score is provided in the

supporting information section (Dataset S1). Interestingly, the

majority of the genes that were highly enriched in our selection

were involved in two major biological processes: LPS core

biosynthesis and regulation of cell shape and morphology. Most

of these candidate genes belong to two long operons (Figure 4B).

In other words, genetic perturbations caused by transposon

insertion in many components of LPS biosynthesis or cell shape

regulation made wild-type cells lose their ability to form a biofilm

in the presence of sPNAG. The dominance of genes involved in

the synthesis and regulation of exposed structural components

suggested that physical interaction between sPNAG and these

surface structures may be a major determinant of biofilm

formation capacity. sPNAG pre-treatment of the cells, however,

did not cause any change in the migration of their extracted LPS

samples on SDS-PAGE gels (Figure S4).

Incomplete disruption of the targeted genes and polar effects are

characteristics of transposon insertion events. In order to get

around these complications and get a fine-scale perspective of

genetic perturbations that prevent cells from responding to

sPNAG, we generated in-frame deletions of some of the candidate

ORFs, obtained from our transposon mutagenesis screen, in the

MG1655 background [23,24]. We then studied the behavior of

these mutant strains in the presence of sPNAG (Table S2). Among

the candidate genes, deletion of rfaY, rafP, or rfaQ, diminished

sPNAG-based biofilm formation (Figure 1, compare i and iii). As

shown in Figure 4C, the product of these three genes are directly

or indirectly involved in addition of phosphate groups to the inner

core of LPS [25].

Since the major common point in the LPS structure of rfaY, rfaP,

and rfaQ mutants is the lower density of negative charge

(phosphate groups) on their LPS outer core, these phosphate

groups are likely to be critical for this interaction. Given the

positive charge of sPNAG, we favor a model in which electrostatic

interaction between the positively charged polysaccharide and the

negatively charged phosphate groups on LPS is the major

determinant of sPNAG-mediated biofilm formation. Electrostatic

interactions were also proposed to be responsible for PNAG-based

biofilm formation in S. epidermidis [20].

It is postulated that neighboring LPS molecules can be cross-

linked by divalent cations due to the presence of phosphate groups

in the LPS structure [25]. Therefore, any presumable electrostatic

interaction between phosphate groups and sPNAG should be

sensitive to increasing concentrations of divalent cations. As shown

in Table S3, response to sPNAG is lost in Ca2+ concentrations

higher than 100 mM, which could be considered as an additional

support for our electrostatic interaction model. However, changing

calcium concentration might also change cell viability. Exposure to

this concentration of calcium, however, did not have any effect on

the viability of the cells, as measured by viable counting and CFU

Figure 3. Schematic representation of the enrichment procedure for transposon mutants defective in sPNAG-based biofilmformation.doi:10.1371/journal.ppat.1000432.g003



determination. Identical results were obtained when the experi-

ment was repeated with other divalent cations (manganese or

magnesium).

Based on the microarray, transposon insertions in genes

involved in regulation of cell shape and morphology should also

interfere with sPNAG-base biofilm formation. However, since all

the genes in this category are essential, we could not introduce

those deletions into the wild-type background and check their

phenotype. Transposon insertion events in this operon presumably

occurred in either regulatory regions (e.g. promoter) or dispensable

parts of essential genes.

As could be inferred from the data, there was no significant

correlation between capacity of producing PNAG and responding

to it. However, in order to confirm this, we studied sPNAG-

induced biofilm formation phenotype of all four DcsrA Dpga

double-mutants (DcsrA pgaA::kan, DcsrA pgaB::kan, DcsrA pgaC::kan,

and DcsrA pgaD::kan) and also all four Dpga single-mutants and

found it to be indistinguishable from that of wild-type cells.

Systematic Characterization of Extra-Genic SuppressorsReverting the Biofilm Formation Defect of DrfaY Cells

We were curious to know whether deletion of rfaY, rfaQ, or rfaP

abolished the response to sPNAG due to downstream signaling

events or the phenotype was a simple consequence of the structural

modifications imposed on LPS. Therefore, we decided to

systematically identify extra-genic suppressors which can restore

biofilm formation capacity of DrfaY cells and test whether there are

any known or putative signaling pathway components among such

suppressors.

To identify suppressors of rfaY deletion, a transposon insertion

library was generated in the DrfaY background (i.e. a strain with

clean deletion of rfaY ORF), and enriched for double mutants that

recovered their ability to form a biofilm in the presence of sPNAG

(opposite to what was demonstrated in Figure 3). A glass slide was

provided as the biofilm formation surface and at the end of each

round, the slide was transferred to a new container with sPNAG

and fresh media. After four rounds of enrichment, macroscopic

microcolony structures could be detected on the glass slide. The

transposon insertion sites in the enriched population were mapped

by the same footprinting strategy described previously (Dataset

S2). Surprisingly, transposon insertions in many of the LPS

biosynthetic genes were significantly enriched in this selection

(Figure 5A)

To validate our microarray predictions, we generated in-frame

deletions of the candidate genes in the DrfaY background and

studied their behavior in the presence of sPNAG (Table S4). We

found that deletion of rfaC, rfaF, rfaI, pgm, galU, rfaH, rfbD, rfbC, and

adiY reverted the phenotype of DrfaY cells. rfaC, rfaF, rfaI, pgm, rfbC,

rfbD, and galU are involved in the synthesis of LPS core structure

or its precursors (Figure 5B). rfbC and rfbD are involved in

rhamnose biosynthesis which is a component of the second major

LPS glycoform in E. coli K12 [26], which is not shown in

Figure 5B. rfaH is a transcription anti-terminator which is required

for full-length transcription of long operons, including rfaQ-K

operon [27]. adiY is the positive regulator of the arginine

decarboxylase system and will be discussed later. Overall, these

mutants make truncated versions of LPS, an expectation we

verified for a subset of them (Figure 5C). We hypothesized that in

these truncated structures, inner phosphate groups of lipid A or

possibly other negatively charged cell-surface moieties (which tend

to be buried by longer LPS chains in the wild-type cells) are now

more exposed and available for interaction with sPNAG. Deleting

any of the four genes in the pga operon did not restore the biofilm

formation capacity of the DrfaY cells, as was also inferred from the

microarray data.

We also used fluorescence microscopy to characterize the

dynamics of biofilm formation in a heterogeneous population

composed of cells either capable (represented by DrfaY DrfaF cells)

or defective (represented by DrfaY cells) in biofilm formation. To

this end, DrfaY cells, expressing RFP fluorescent marker, were

competed against DrfaY DrfaF cells expressing GFP with the

starting ratio of 1:1 for making a biofilm on a glass slide in the

presence of sPNAG. After 12 hours, the biofilm structure formed

on the glass slide was visualized by fluorescence microscopy. As

shown in Figure 5D, top row, microcolonies in biofilm structure

were mostly formed by DrfaY DrfaF double mutants (i.e. GFP

expressing cells). The same result was obtained by swapping the

fluorescent labels (Figure 5D, bottom row).

Transposon insertions typically lead to a loss of function

phenotype. In order to complement our transposon insertion based

approach, we used an over-expression library in the DrfaY

background. This library contained ,2.56105 independent mutants

each carrying a 1–3 kb long genomic fragment of E. coli cloned into

the pBR322 plasmid. The over-expression library was enriched for

mutants responding to sPNAG, similar to the approach used for

studying the DrfaY transposon insertion library. After four rounds of

enrichment, the over-expressed fragments represented in the

enriched population were identified by microarray hybridization

(Dataset S3). As expected, when comparing the results from DrfaY

transposon insertion library with DrfaY over-expression library, LPS

biosynthesis genes showed the opposite behavior (Figure 6A). Many

LPS biosynthetic genes like rfaG, rfaJ, rfaI, rfaP, kdtA, rfbX, rfaZ, and

rfaL were found to be among the top 10% highly enriched category

in the DrfaY background transposon insertion library while in the

over-expression library, they belonged to the top 10% most depleted

group.

The microarray results also revealed that mutants over-

expressing certain genes associated with the acid tolerance system

in E. coli were abundant in the enriched population. In order to

clarify how this system contributes to biofilm formation, we further

characterized the phenotypic consequences of over-expressing

these genes in individual cells. To this end, we isolated individual

Figure 4. Genome-wide identification of loci involved in sPNAG-based biofilm formation. (A) Distribution of z-scores after enrichment formutants impaired in sPNAG-based biofilm formation. A transposon insertion mutant library of E. coli has been enriched for mutants that are defectivein responding to sPNAG. To quantify the contribution of different loci to this phenotype, the insertion sites of the transposon in the enrichedpopulation were mapped using a microarray-based approach. The average signal acquired for each ORF on the microarray from two replicateexperiments was used to calculate the z-score value for that ORF. This value reflects the relative abundance of transposon insertion events in the ORF(or in its vicinity) in the enriched population compared to the maximally diverse parental library. Distribution of these z-scores in the enrichedpopulation is illustrated by a histogram. Some of the ORFs which were significantly enriched were labeled on the histogram. More detailedinformation regarding the calculation of z-score is provided in Dataset S1. (B) Many of the transposon insertions with high z-scores in the firstselection belonged to one of the two long gene clusters involved in LPS biosynthesis (top row) or regulation of cell shape and peptidoglycanbiosynthesis (bottom row). ORFs with high z-scores in the selection are shown in red. (C) Structure of the major glycoform of E. coli K-12 LPS [26]together with its biosynthetic genes (enzymes). The three genes (enzymes) whose deletion abolished sPNAG-based biofilm formation are highlightedin red.doi:10.1371/journal.ppat.1000432.g004



clones from the enriched population. One of the isolated mutants

was found to have the genomic region corresponding to three

genes, gadW, gadY, and gadX. GadX and GadW are dual regulators

of the glutamate-dependent decarboxylase acid-resistance system

of E. coli. [28]. GadY is a small RNA which acts as a positive

regulator of gadX [29].

Figure 5. Characterization of extra-genic suppressors reverting the biofilm formation defect of DrfaY cells in a transposonmutagenized library. (A) Distribution of z-scores after enrichment for DrfaY double mutants that recovered the capacity for sPNAG-based biofilmformation. (B) Schematic representation of LPS structure, together with its biosynthesis genes (enzymes) in DrfaY cells. Secondary mutations whichreverted the biofilm-formation deficiency of the DrfaY cells are highlighted in red. (C) LPS samples from a subset of DrfaY double mutants thatrecovered their ability to respond to sPNAG are separated on a SDS-PAGE gel. All double mutants showed truncated versions of LPS compared to theDrfaY cells. LPS samples from DrfaY DrfaC and DrfaY DrfaF cells were not detectible on the gel. (D) DrfaY cells, expressing RFP (mCherry), and DrfaYDrfaF cells, expressing GFP, were competed against each other for biofilm formation on a glass slide surface in presence of sPNAG (top row). Threeimages from left to right show the red channel, green channel, and the merged version. Similar results were obtained after swapping the fluorescentmarkers (bottom row).doi:10.1371/journal.ppat.1000432.g005



Our previous results demonstrated that LPS modification was a

dominant mechanism in regulating response to sPNAG. There-

fore, we first investigated the effect of gadXYW over-expression on

LPS structure. As shown in Figure 6B, DrfaY pBR322-gadWYX

strain contained some smaller LPS variants as compared to the

parental DrfaY cell. This suggests that the reversion of DrfaY

phenotype upon over-expression of gadWYX gene cluster is a

consequence of this truncated LPS structure. To test whether this

change in LPS structure was due to a transcriptional regulatory

event, we measured the transcription of the rfaQ-K operon in both

pBR322-gadWYX and pBR322 (empty vector) backgrounds by a b-

galactosidase assay and found it to be almost 3-fold lower in the

gad-over-expressing cells (Figure 6C). Evidence regarding the

existence of a cross-talk between acid tolerance system and LPS

regulation has been observed before, and gadE, the transcriptional

regulator of the acid resistance system in E. coli, was reported to be

a potential activator of the rfaQ-K operon [30]. We also found that

over-expression of gadY alone was sufficient to cause the

phenotype, although not as strongly as the over-expression of

gadXYW.

A strain harboring the pBR322-argR plasmid was also isolated

from the enriched DrfaY library. However ArgR, the negative

regulator of arginine biosynthesis system, acts as a weak suppressor

of DrfaY biofilm formation deficiency. Putting all these observa-

Figure 6. Characterization of extra-genic suppressors reverting the biofilm formation defect of DrfaY cells in an over-expressionlibrary. (A) Distribution of LPS biosynthesis genes were compared in the DrfaY transposon insertion and over-expression libraries. The microarrayoutputs of the DrfaY transposon insertion and the DrfaY over-expression libraries were sorted separately based on their z-score and divided into 10equally populated bins. The number of genes belonging to lipopolysaccharide biosynthesis gene cluster (GO index GO:0009103) in each bin wascounted and used to calculate hypergeometric p-value for over-representation of LPS biosynthesis genes in that bin for each library. Each bin wascolor-coded based on its 2log10(p-value). The yellow color for a bin reflects statistically significant abundance of lipopolysaccharide biosynthesisgenes in that bin. As shown, many LPS biosynthetic genes were found to be among the top 10% highly enriched category in the DrfaY backgroundtransposon insertion library, while in the over-expression library they belonged to the top 10% most depleted group. (B) LPS samples extracted fromDrfaY pBR322-gadXYW and DrfaY cells and separated on SDS gels. (C) Transcription level of the rfaQ-K operon promoter is compared between DrfaYpBR322-gadWYX and DrfaY pBR322 cells by a b-galactosidase assay.doi:10.1371/journal.ppat.1000432.g006



tions together, four of the suppressors found in the transposon

insertion and over-expression libraries, adiY, gadX, gadW, and argR,

were directly or indirectly associated with the amino acid

decarboxylase systems, involved in acid tolerance in E. coli. The

biological function of these suppressors (Table S5) and the

distribution of acid tolerance genes in the over-expression library

(Figure S5) suggest that down-regulation of the acid stress

response, or more specifically the amino acid decarboxylase

systems, positively contribute to biofilm formation in the DrfaY

background, presumably due to the changes imposed on LPS.

A Model for sPNAG-Mediated Biofilm Formation: FromPassive Nucleation to Active Maturation

Overall, our observations support the existence of a physical

interaction between sPNAG and LPS. As such, biofilm formation in

the presence of sPNAG may be a purely structural phenomenon,

occurring as a simple consequence of passive interactions between

sPNAG and LPS. Based on this model, even dead cells with intact

outer membrane structure should still be capable of responding to

sPNAG. To test this, we killed wild-type cells by either UV-

irradiation or exposing them to formaldehyde, and visualized their

behavior upon exposure to sPNAG. Time-lapse microscopy showed

that these dead cells start nucleating microcolony structures, similar

to living cells (Video S3). All together, our observations argue that

sPNAG-mediated biofilm formation can be considered as a two-step

process, starting with the nucleation event which is a purely

structural phenomenon, followed by microcolony expansion and

maturation which is a growth-dependent process (Figure 1, compare

iv and vii to v and vi).

From Laboratory Strains to Clinical IsolatesMost natural and clinical isolates of E. coli produce different

serotype-specific surface structures including O-antigen and

capsular polysaccharide, also known as K-antigen, which are

absent in E. coli MG1655 [31,32]. We were curious to know how

variations in composition of these surface antigens might affect the

response to sPNAG. Therefore, we chose 11 strains, which were

reported to be competent of endogenous PNAG production [8],

for further analysis. Among these strains, 7 formed biofilms in the

presence of sPNAG (Table S6), and two of the latter group were

also O-antigen2 (Figure S6). In case of K-antigen, we focused on

K1 capsule, a homopolymer of a-(2-8)-linked polysialic acid [33],

which is the predominant capsule found in a major subset of these

clinical isolates. Three K1+ isolates used in this study were also

capable of responding to sPNAG (Table S6).

Natural and clinical isolates of E. coli possess uncharacterized

surface structures other than K1, e.g. fimbriae and other capsular

polysaccharides. These structures could significantly affect the

physicochemical properties of the cell surface. Consequently, cell

response to sPNAG in these clinical isolates could not be solely

judged based on their O-antigen structure or presence of K1

capsule. Furthermore, the limited number of strains tested in this

study and their non-isogenic background make these observations

preliminary and they should be followed up with future studies.

Therefore, we decided to study the role of O- and K-antigen in the

well-characterized K-12 background.

We generated all possible combinations for presence or absence

of O16 antigen and K1/K92 capsule in the K-12 background.

K92 is a polysialic acid capsule very similar to K1, and its

biosynthetic gene cluster can be transferred on a plasmid. As

shown in Table 1, only non-capsulated O16+ cells were impaired

in sPNAG-based biofilm formation. In the presence of both O16

and K1/K92 antigen, however, cells were capable of responding

to sPNAG, which is not surprising considering that capsule is a

more exposed surface structure than O-antigen [32]. Since K1

and K92 capsules confer a high density of negative charge to the

E. coli cell surface, their presence could contribute to establishing

any potential electrostatic interaction with sPNAG. These data

suggest that loss of O-antigen (O16) or presence of K1 capsule is

associated with sPNAG-induced biofilm formation in the E. coli K-

12. However, in order to confirm the involvement of these

structures in sPNAG-induced biofilm formation, targeted genetic

experiments together with more careful characterization of their

role in physiochemical properties of outer membrane are required.

sPNAG-Based Biofilms: What Do Biofilms Share inCommon?

Biofilms can afford protection from a variety of environmental

challenges including phagocytosis, extreme pH, and antibiotic

exposure [1]. We were curious to characterize some of these

biofilm-specific features in sPNAG-based biofilms. In order to

investigate their antibiotic tolerance, we challenged the cells with

two different antibiotics: ampicillin and polymyxin B. As shown in

Figure 7, cells in the context of sPNAG biofilm showed ,10 fold

higher tolerance to polymyxin B compared to planktonic cells,

whereas no significant difference was observed in tolerance to

ampicillin. Polymyxin B resembles antimicrobial peptides, an

integral component of the innate defense system in many

organisms, in terms of both structure and mechanism of action

[34]. Considering sPNAG as a positively charged matrix encasing

the cells, electrostatic repulsion between this polysaccharide and

polymyxin B could potentially protect the bacteria by reducing the

local concentration of the drug in the vicinity of the cells.

However, we still consider the involvement of other as yet

unknown sPNAG-induced physiological response in this phenom-

enon.

Table 1. Role of surface antigens in sPNAG-based biofilm formation.

Presence (+) or Absence (2) of O (O16) or K (K1/K92) Antigen

O-Antigen 2 +(O16) 2 2 +(O16) +(O16)

K-Antigen 2 2 +(K1) +(K92) +(K1) +(K92)

Response to sPNAG + 2 + + + +

O-antigen production was restored in MG1655 by plasmid pMF19, encoding a functional copy of rhamnosyl-transferase, RfaL, whose gene is mutated in E. coli K-12 [31].Transformation of MG1655 strain with this plasmid allowed production of O16 antigen [45] confirmed on SDS-PAGE gels (Figure S6, compare lane 8 and 9). For K-antigen, only K1 and K92 capsules were considered. In the case of the K92 capsule, the gene cluster responsible for its biosynthesis was cloned into the pGB20 plasmidand could be easily transferred to any background [46]. As a K1+ cell, a K-12-based K-12/K1 hybrid (EV36), generated by conjugation, was used which was capable ofproducing a polysialic acid capsule indistinguishable from that of natural K1 strains [47]. All the experiments were performed in the MG1655 background except forthose corresponding to O162K1+ and O16+K1+ strains, which were in the EV36 background.doi:10.1371/journal.ppat.1000432.t001



Considering other biofilm-specific phenotypes, we found no

significant change in the rate of F-plasmid conjugation in sPNAG-

based biofilms (Figure S7). We also did not observe any evidence

supporting the existence of a phase-variable mechanism regulating

sPNAG production in E. coli (Figure S8 and Figure S9).

We were also curious whether sPNAG can induce biofilm

formation in species other than E. coli, because this could

potentially facilitate interspecies interactions, e.g. conjugation,

between E. coli and other microorganisms. Therefore, we exposed

Salmonella typhimurium LT2 cells to sPNAG similar to what was

done for E. coli, but we did not observe any detectable biofilm

formation. We reasoned that if sPNAG-mediated biofilm forma-

tion has a structural basis, then a mutant Salmonella with modified

surface characteristics may be capable of forming a biofilm in the

presence of sPNAG. To test this hypothesis, we applied sPNAG to

a transposon insertion library of S. typhimurium LT2 with

approximately 105 mutants and enriched for mutants that form

biofilms. After four rounds of enrichment, macroscopic microcol-

ony structures were formed by cells. We mapped the transposon

insertion sites in 4 of the mutants capable of forming biofilms in a

sPNAG-dependent pattern. In all cases, the transposon mapped to

different positions in the rfaK gene, involved in LPS biosynthesis.

Salmonella rfaK mutants are lacking most of the O-antigen structure

[35], providing further support, beyond E. coli, for our physical

interaction model between sPNAG and LPS.

Discussion

Poly-N-acetylglucosamine, the major virulence factor of Staph-

ylococcus epidermidis, has recently been found in many other

pathogenic bacteria [7], including E. coli, but PNAG-based

biofilms in these pathogens are poorly characterized relative to

Staphylococcus species [2]. Here, we carried out a systematic genetic

analysis of poly-N-acetylglucosamine-induced biofilm formation in

E. coli. However, instead of working in a biofilm-permissive genetic

background, in which the time scale of biofilm formation is slow,

we applied the functionally active secreted version of PNAG

(sPNAG) to wild-type E. coli MG1655 cells and observed rapid and

reproducible biofilm formation. The fast kinetics and robustness of

sPNAG-induced biofilm formation phenomenon allowed us to

comprehensively characterize its underlying genetic basis. Our

observations support the notion that electrostatic interaction

between positively charged sPNAG and different cell surface

antigens with negative charge is responsible for the formation of

the biofilm structure. This is consistent with the generally accepted

intuition that physicochemical properties of the matrix, including

its charge, geometry, ion selectivity, and pore size contribute

significantly to biofilm formation [1,18].

The composition and physicochemical properties of cell surface

structures can be modulated by multiple biological pathways and

environmental factors. Therefore, although response to sPNAG

seems to be the consequence of simple electrostatic interactions, it

can be regulated by a complex interplay between LPS structural

dynamics, the presence of serotype-specific capsular polysaccha-

ride, the acid tolerance system, and cell morphology as shown in

this work. There are also other relevant pathways, not fully active

in our laboratory strain, that might contribute to this phenotype,

such as addition of positively charged 4-amino-4-deoxy-L-

arabinose to lipid A involved in polymyxin B and other cationic

antimicrobial peptide tolerance in E. coli and S. typhimurium.

Interestingly, a polymyxin B resistant mutant isolated from our

transposon insertion library was also impaired in responding to

sPNAG (data not shown). Since resistance to polymyxin is usually

concomitant with higher density of positive charge on the outer

membrane [36], polymyxin resistant mutants are expected to be

defective in interacting with sPNAG. Given that antimicrobial

peptides generated by the host immune system are a major

challenge for the survival of pathogens, encasement in a positively

charged matrix or biofilm serves as a protective mechanism

against antimicrobial peptides for sensitive cells, while resistant

cells can survive without it.

Since in this study, selections were performed with pools of

mutants rather than with clonal populations, phenotype of

different mutants should be interpreted as a spectrum of different

capacities for forming a biofilm rather than a simple binary

phenomenon. The results of our selection, carried out on the over-

expression library in the DrfaY background, strongly support

this notion. DrfaY cells are defective in responding to sPNAG, so it

is reasonable to assume that only a handful of genes in the

Figure 7. Antibiotic tolerance in sPNAG-based biofilms. 108 stationary phase wild-type cells were used to inoculate fresh LB in the presence(first and third column from left) or absence (second and fourth column) of 0.1 U/ml of sPNAG. After 12 hours, polymyxin B or ampicillin was addedto a final concentration of 10 mg/ml and 100 mg/ml, respectively. After 18 hours, cells were pelleted down, washed with PBS three times, treated withDispersin B to break the biofilm structures (if any), and plated for CFU determination. Independent experiments showed that Dispersin B treatmentdid not change the viability of the cells. For each column, results from ten independent samples were averaged and reported. Error bars indicate thestandard error of the mean calculated from all independent measurements.doi:10.1371/journal.ppat.1000432.g007



over-expression library should be capable of suppressing its

phenotype and the majority of the library mutants should be

equally defective in biofilm formation. However, the fact that a

considerable fraction of genes involved in LPS biosynthesis were

significantly depleted in that selection (Figure 6A) indicates that

even DrfaY cells might have residual biofilm formation activity and

that over-expression of some LPS biosynthetic genes might reduce

this partial activity. The weak biofilm formation activity of DrfaY

cells could be due to their entrapment in the microcolony

structures formed by other mutants in the population. This is in

part shown in Figure 5D in which DrfaY cells are mostly

colocalized with microcolonies rather than being uniformly

distributed. Imposed modifications in LPS structures caused by

over-expression of some LPS biosynthetic genes [37] might

interfere with this partial activity.

In this study, we have demonstrated the involvement of different

surface structures and regulatory systems in sPNAG-mediated

biofilm formation. However, there might exist a myriad of other

uncharacterized regulatory systems, surface structures, and

environmental factors that may influence this phenotype through

changes in the physicochemical properties of the cell surface.

Covalent modifications of lipid A with phosphoethanolamine in a

Ca2+-dependent manner, or with 4-amino-4-deoxy-L-arabinose in

response to Mg2+ and pH, represent just a few examples [38].

Therefore, even if a natural isolate of E. coli is incapable of

responding to sPNAG in the laboratory environment, it may show

a different phenotype in its natural environment. There is also the

possibility of higher level cooperation between different cell types

in the population, with one subpopulation producing sPNAG

while another is responding to it. The producer subpopulation

may even be defective in the initiation of the biofilm formation

process, but could be incorporated into preformed structures.

Examples of population heterogeneity in biofilm communities has

been reported before [1]. Furthermore, it is likely that different

cells produce different versions of the PNAG polymer, with

different degrees of acetylation [39] or different polymer length

distributions, which could make response to sPNAG more strain-

specific. A better understanding of sPNAG-based biofilms

acquired from studies like this could be useful for development

of new therapeutic strategies against pathogens that use this

polysaccharide as a virulence factor. The systematic framework

presented in this study, along with the acquired insights, should

also benefit the study of microbial biofilms formed by other

species.

Materials and Methods

Strains, Media, and Microbiological TechniquesAll strains used in this study are listed in Table S7,

bacteriophages and plasmids are mentioned in Table S8. All the

experiments were performed in LB (1% tryptone, 0.5% yeast

extract, 1% NaCl), supplemented as required with the following

antibiotics: ampicillin, 50 mg/ml; tetracycline, 25 mg/ml; specti-

nomycin, 100 mg/ml; chloramphenicol, 30 mg/ml and kanamy-

cin, 100 mg/ml, unless otherwise mentioned. b-galactosidase

measurements were performed in triplicate as described before

[40].

DNA ManipulationsTransposon mutagenesis and microarray based genetic foot-

printing were carried out as described before [22]. Chromosomal

deletions were created using the previously described method [23]

and transferred by generalized transduction with P1 phage as

required. lacZ reporter strain was generated using the plasmid

pCE37 [41]. The over-expression plasmid library construct was a

kind gift of Joseph Sklar. Each plasmid contained a 1–3 kb long

fragment of E. coli MC4100 genome (average fragment size of

2 kb) cloned into the b-lactamase coding sequence of the pBR322

vector. The plasmid pool was electroporated into E. coli MG1655

DrfaY cells, leading to ,2.56105 independent over-expressing

mutants. For PCR amplification of DNA constructs for cloning,

Pfu Ultra polymerase (http://www.stratagene.com) was used,

whereas ExTaq DNA polymerase (http://www.takara-bio.com)

was used for all other PCR reactions. Restriction endonucleases

and T4 ligase were obtained from New England Biolabs (http://

www.neb.com). DNA purification kits were obtained from

QIAGEN (http://www1.qiagen.com). Primer sequences are

available upon request.

LPS Extraction and AnalysisStructural analysis of LPS samples was performed as described

before [42]. LPS was purified from cell lysates after phenol-ether

extraction, separated on 14% tricine-SDS-PAGE gel and

visualized after silver staining.

MicroscopyAll fluorescence and light microscopy experiments were

performed using Zeiss AxioVision 4.5. Time-lapse microscopy

was performed in the FC81 flow cell apparatus from Biosurface

Technologies Corp (in the absence of flow). SEM analysis was

performed using a Philips XL30 Field Emission Scanning Electron

Microscope.

Determination of Hexosamine Content in the SpentMedia

For the sake of consistency, a large batch of DcsrA spent media,

grown in LB, was prepared and used as the source of sPNAG for

all the experiments throughout this study. The amount of sPNAG

present in this stock solution, was estimated by measuring its

hexosamine content using 3-methyl-2-benzothiazolone hydrazone

hydrochloride (MBTH) method [43] and found to be ,0.175 mg

equivalent of hexosamine per ml of saturated DcsrA culture

supernatant. The quantity of sPNAG in each milliliter of this stock

solution was defined to be 1 arbitrary unit (U). Control

experiments were also carried out using sPNAG from different

preparations of DcsrA spent media and the results were

reproducible.

sPNAG Purification and Mass SpectrometrySpent media from 1 liter of saturated DcsrA culture in LB was

passed through a 0.22 m filter (http://www.nalgene.com/). The

cell-free supernatant was concentrated 200 fold by Centriplus YM-

100 columns (http://www.millipore.com). The concentrated

sample was treated with DNase I (2 mg), RNase A (10 mg), and

a-amylase (20 mg) and incubated for 2 hours at room temperature

followed by 2 hours at 37uC. Next, sample was treated with

Proteinase K (20 mg) for 1 hour at 37uC and 1 hour at 55uC.

Enzymes and other proteins were removed by pre-warmed (55uC)

phenol: ether extraction, followed by ethanol precipitation. The

precipitate was re-suspended in water and fractionated on a S-300

Sephacryl column. The fractions with biofilm-inducing activity

were pooled together and concentrated by Centriplus YM-100

columns. The sample was treated with Dispersin B (20 mg) for

4 hours at 37uC. After removing the enzyme by phenol:

chloroform extraction, the sample was dialyzed with Spectra/

Por cellulose ester membranes with MWCO = 500 (http://www.

SPECTRUMLABS.com/) to remove the salt present in Dispersin



buffer. Finally, the sample was analyzed by an ESI-LTQ Orbitrap

Hybrid mass spectrometer from Thermo Fisher Scientific.

sPNAG-Induced Biofilm Formation AssayFor all biofilm formation assays, cells and sPNAG were added to

fresh LB in such a way as to obtain ,56108 cells and 0.1 U of

sPNAG per milliliter of the mixture. Biofilm structures were

studied or transferred (in case of serial enrichments) 12 hours after

exposure to sPNAG at 25uC.

Preparation of Over-Expression Library Sample forMicroarray Hybridization

Plasmids from ,109 cells of the enriched population (or in case

of references, from the maximally diverse unselected library) were

extracted using QIAGEN plasmid miniprep kit. The extracted

plasmid pool was amplified with two separate PCR reactions,

using primer pair pBR_Lib_T7_L and pBR_Lib_R or

pBR_Lib_T7_R and pBR_Lib_L. The sequences of these primers

are as follows:

pBR_Lib_T7_L: 59-GTCAACCTGGCTTATC-

GAAATTAATACGACTCACTATAGGGCTCT-

TACTGTCATGCCATCCGTA-39

p B R _ L i b _ R : 5 9 - G T T T T C C A A T G A T G A G -

CACTTTTA-39

pBR_Lib_T7_R: 59-GTCAACCTGGCTTATC-

GAAATTAATACGACTCACTATAGGGGTTTTC-

CAATGATGAGCACTTTTA-39

pBR_Lib_L: 59-ATAATTCTCTTACTGTCATGC-

CATCCGTA-39

The incorporated T7 promoter in pBR_Lib_T7_L and

pBR_Lib_T7_R is underlined. Cycling conditions for PCR were

94uC for 2 min; 30 cycles of 94uC for 30 s, 55uC for 30 s, and

72uC for 5 min; and 72uC for 10 min, using ExTaq DNA

polymerase. The PCR products from these two reactions were

pooled together. The T7 promoter incorporated into the

pBR_Lib_T7_L and pBR_Lib_T7_R primer sequences was used

to generate RNA from the pooled PCR product in an in vitro

transcription reaction using Ambion MEGAscript T7 Kit (http://

www.ambion.com). Finally the RNA from the previous reaction

was reverse transcribed to cDNA, using Cy3-labeled nucleotides

with Invitorgen SupreScript II Reverse Transcriptase (http://

www.invitrogen.com). The fluorescently labeled cDNA was used

for microarray hybridization versus Cy5-labeled fragmented

(nebulized) MG1655 genomic DNA. The hybridization data of

the two maximally diverse unselected libraries were used as the

reference.

Mapping Spontaneous Mutations Abolishing PNAGProduction

Production of PNAG in S. epidermidis could be discriminated on

congo red indicator plates [44]. In order to extend this to E. coli,

congo red indicator plates (3% tryptic soy broth, 1% glucose,

0.08% congo red, and 1.5% agar) were supplemented with

200 mM NaCl which was reported to enhance pga locus

transcription in E. coli [16]. Different dilutions of stationary phase

cultures of DcsrA mutants were plated on these plates and

incubated for two days at 37uC. PNAG-producing DcsrA cells

formed dark brown colonies whereas DcsrA mutants defective in

producing PNAG were red. To map the location of the mutation

which abolished PNAG production, red colonies were picked

individually and transduced with a P1 phage lysate obtained from

the maximally diverse transposon insertion library. Kanamycin

resistant transductants which recovered their PNAG production

were identified by replica plating on congo red plates. Dark

colonies were picked, and the location of transposon insertion,

which should have been linked to the mutation site, was mapped

using the same footprinting strategy. The exact location of the

mutation was determined after PCR amplification of the candidate

genomic locations and subsequent sequencing of the PCR

product.

Supporting Information

Dataset S1 Microarray data for identification of transposon

insertion mutants defective in responding to sPNAG. The z-score

is calculated based on the average microarray signal of two

independent replicates normalized versus five hybridizations of the

maximally diverse unselected library. First, the average hybrid-

ization intensity value of every ORF from duplicate selection

experiments (x) is normalized using mean (,x.) and standard

deviation (std(x)) of five hybridizations of the unselected library,

representing the null distribution for that ORF. The z-score is

calculated using the following equation: z = (x2,x.)/std(x).

Therefore, the z-score value for each ORF reflects the relative

abundance of transposon insertion events within that ORF or in its

vicinity in the enriched population versus the unselected library.

More detailed information on hybridization values of each

replicate and unselected library is provided in a separate

worksheet.

Found at: doi:10.1371/journal.ppat.1000432.s001 (1.78 MB XLS)

Dataset S2 Microarray data for identification of transposon

insertion mutants suppressing rfaY deletion phenotype. The z-score

is calculated based on the average microarray signal of two

independent replicates as explained before (Dataset S1). More

detailed information on hybridization values of each replicate and

unselected library is provided in a separate worksheet.


Dataset S3 Microarray data for identification of over-expressed

fragments suppressing rfaY deletion phenotype. The normalized

score is calculated by subtraction of the average microarray signal

of two maximally diverse unselected libraries from that of two

experimental replicates. More detailed information on hybridiza-

tion values of each replicate and unselected library is provided in a

separate worksheet.


Figure S1 Reporter assay for pga operon activity in the presence

or absence of csrA deletion. Up-regulation of the pga locus

transcription in the DcsrA background was confirmed by a reporter

assay. pga promoter was placed upstream of a GFP-coding

sequence on a multi-copy plasmid (pPGA’-GFP). The reporter

plasmid was electroporated into both wild-type and DcsrA strains.

As shown here, DcsrA cells strongly express gfp while wild-type cells

do not show detectable fluorescence. Scale bars, shown in red,

correspond to 2 mm.

Found at: doi:10.1371/journal.ppat.1000432.s004 (0.09 MB PDF)

Figure S2 Comparative ESI LTQ OrbiTrap mass spectrum of

Dispersin B-digested PNAG isolated from E. coli and S. aureus,

acquired in positive mode. PNAG sample from E. coli was isolated

and purified as described in the Materials and Methods section. S.

aureus PNAG which was isolated from strain MN8m ([1] in Text

S1) was a kind gift from Gerald Pier. Purified polysaccharide

samples were treated individually with Dispersin B for 1 hour at

37uC and passed through Centriplus YM-10 columns. The flow-

through was analyzed by an ESI-LTQ Orbitrap Hybrid mass



spectrometer from Thermo Fisher Scientific. The mass spectrums

show abundance of different molecular ions (characterized by their

m/z value) relative to the most abundant ion. As shown here (E.

coli, panel A, and S. aureus, panel B), there are many peaks with

identical m/z values in both spectra, indicating that the isolated

polysaccharides are closely related polymers. A complete list of m/

z values for all potential mono- and oligosaccharides species that

could be generated from an incomplete digestion of a PNAG

sample with all possible acetylation patterns are also given in

Table S1, as a reference. Panel C shows the mass spectrum of

undigested E. coli PNAG and panel D corresponds to the spectrum

of Dispersin B enzyme. As shown in panel C, intact PNAG can not

be analyzed efficiently by OrbiTrap mass spectrum due to its large

molecular weight. Therefore, the intensity of peaks acquired for

this sample is lower than the noise (e.g., compare the relative

intensity of the background peak with m/z value of 131.00 in

panels C and D). Since digested PNAG samples in panels A and B

were passed through a YM-10 filter, those samples should be free

from Dispersin B, and spectrum of the enzyme (panel D) was only

provided as a control. References in all supporting figures and

tables can be found in Text S1.


Figure S3 SEM images of wild-type cells treated with sPNAG

(400,0006, 200,0006, and 100,0006, respectively). 50 ml of fresh

LB was inoculated with ,2.561010 wild-type MG1655 cells with a

sPNAG content of 0.1 U per milliliter of the mixture at 25uC. A

glass slide was provided as biofilm formation surface in a vertical

orientation. The biofilm formed on the surface of the glass slide

after 12 hours was fixed with 2.5% glutaraldehyde in 200 mM

sodium cadodylate for 60 minutes. Next, it was gently washed

twice with 200 mM sodium cadodylate. The sample was post-fixed

with 1% osmium tetroxide in sodium cacodylate buffer for

30 minutes. It was washed 4 times, 10 minutes each, with distilled

water to remove all the fixative and buffer salts. Next, it was

dehydrated sequentially in 35%, 45%, 55%, 65%, 70%, 85%,

95%, and 100% ethanol, 5 minutes each. Then, it was transferred

first to 50% ethanol: 50% TMS (tetramethylsilane) followed by

20% ethanol: 80% TMS, 15 minutes each. All the above-

mentioned steps were carried out at 4uC. The sample was allowed

to air dry at room temperature, and coated with palladium/gold.

Finally, the biofilm structures were visualized by a Philips XL30

Field Emission Scanning Electron Microscope.


Figure S4 SDS-PAGE-based analysis of sPNAG-LPS interac-

tion. LPS samples were extracted from wild-type and DrfaY cells

previously treated with sPNAG (1 U/ml) for 12 hours or nothing

(as control). The samples were separated on a 14% SDS gel and

silver stained. As shown here, sPNAG pre-treatment of the cells

did not cause any change in the migration of their LPS samples on

SDS-PAGE gel. This could be due to multiple reasons, including

the large molecular weight of sPNAG, which is out of the

resolution-range of the gel, presence of SDS in the gel and buffer

which interferes with electrostatic interactions, or requirement for

other components for establishment of the interactions which are

present in vivo. These results do not rule out the possibility of

electrostatic interactions between sPNAG and LPS.


Figure S5 Distribution of acid stress genes in DrfaY over-

expression library. Distribution of acid stress response genes in the

microarray result of the DrfaY over-expression library. Sorted

microarray data is divided into 10 equally populated bins and

number of genes belonging to ‘‘Acid Stress Response Up-

regulated’’ category in each bin was counted and used to calculate

hypergeometric p-value to represent the statistical significance of

over-representation of those genes in that bin. The bins are

pseudo-colored based on the 2log10(p-value), with yellow color

indicating over-representation. Generation of a similar plot has

been explained in more detail in Figure 6A.


Figure S6 O-antigen profiling of clinical and laboratory strains

of E. coli. LPS samples extracted from 11 clinical isolates of E. coli

together with two samples from the MG1655 strain with or

without plasmid pMF19 are analyzed on SDS gels after silver

staining. Plasmid pMF19 contains a functional copy of rhamnosyl-

transferase gene, rfaL, which is mutated in E. coli K-12 ([2] in Text

S1). Transformation of MG1655 strain with this plasmid allows

production of O16 antigen ([3] in Text S1). Among the 11 clinical

strains, there were 7 urinary tract infection (UTI) isolates,

including UTI-E, -G, -H, -J, -P, -R, and -U, and 4 blood isolates

from patients in neonatal intensive care units (NICU), including

NICU-2, -4, -10, and -12 ([4] in Text S1). Based on the gel, UTI-

U was clearly O-antigen2 and UTI-P showed only the first smooth

LPS band together with the lipid A-core band. Mutants with LPS

banding pattern similar to UTI-P were reported not to react with

anti-O antiserum and are considered phenotypically O-antigen2

([5] in Text S1). UTI-E and UTI-G produced less of high

molecular weight O-antigen variants, but they were clearly O-

antigen+ when the gel was overloaded (data not shown).


Figure S7 Horizontal gene transfer rate in sPNAG-based

biofilms. Some microbial biofilms promote the rate of horizontal

gene transfer, facilitating the spread of antibiotic resistance genes

([6] in Text S1). An equal number (,108) of donor (tetracycline

resistant) and recipient (chloramphenicol resistant) E. coli cells were

mixed together in LB in the presence or absence of 0.1 U/ml of

sPNAG. After 12 hours, serial dilutions of each sample were

plated on LB+tetracycline+chloramphenicol plates for CFU

counting of conjugants. For each case, results from ten

independent samples were averaged and reported. Error bars

indicate the standard error of the mean calculated from all

measurements. As shown, no significant correlation exists between

the presence/absence of sPNAG-based biofilm and the rate of

horizontal gene transfer.


Figure S8 Congo red indicator plates for discrimination of

sPNAG production in E. coli. sPNAG-producing DcsrA cells (dark

brown colonies, left) can be discriminated from otherwise isogenic

DcsrA mutants defective in producing sPNAG (red colonies, right)

based on their colony color. Production of PNAG in S. epidermidis

can be distinguished on congo red indicator plates ([7] in Text S1).

In order to extend this to E. coli, congo red indicator plates were

supplemented with salt, which was reported to enhance pga locus

transcription in E. coli ([8] in Text S1). As shown here, on the

indicator plate, sPNAG-producing DcsrA cells formed dark brown

colonies whereas DcsrA pgaA::kan cells (which had lost their ability

to produce sPNAG) were red.


Figure S9 Identification of mutations that abolished sPNAG

production in the DcsrA background. In S. epidermidis, PNAG

production is subject to a phase-variable regulatory mechanism

([7] in Text S1). In order to understand whether production of

sPNAG in E. coli undergoes phase variation as well, three mutants

were isolated from independent DcsrA cultures which had lost their

sPNAG production ability based on their colony color on congo

red indicator plates (Figure S8). When transformed with the



plasmid pPGA’-GFP, two of them could express reporter gfp

downstream of the pga promoter while the third one could not.

Using generalized phage transduction, the genomic location of the

mutation in these three mutants was mapped. Upon sequencing

the candidate genomic locations, the identity of the mutations

were found. One mutant (mutant I) had an insertion element

(IS1E) in 302nd nucleotide of pgaC ORF. In the second mutant

(mutant II), the whole divergent intergenic region between pga

operon and ycdT gene together with the first 611 nucleotides of

ycdT ORF and first 1009 nucleotides of pgaA ORF was substituted

by IS1E. The third mutant (mutant III) was found to have a

deletion spanning the entire nhaR ORF together with 28 upstream

and 176 downstream nucleotides. NhaR is required for PNAG

production in E. coli as it activates pga operon transcription ([8] in

Text S1). This explains why mutant III does not fluoresce when

transformed by the reporter plasmid pPGA’-GFP. In S. epidermidis,

phase variation in PNAG production is mostly controlled by

insertion and excision of an insertion sequence element in PNAG

biosynthetic genes ([7] in Text S1), similar to what is happening in

case of mutant I and II. However, no reversion back to the

producing state was observed in any of these three mutants. These

data suggest that inactivation of PNAG production in the DcsrA

background is presumably due to spontaneous loss of function

mutations rather than a programmed phase variation process. The

high level of PNAG production in the DcsrA background imposes a

considerable energy burden on the cell, therefore loss of function

mutations in PNAG biosynthetic pathway may be strongly selected

for in this background.


Table S1 Expected m/z values for all possible Dispersin B

digested PNAG fragments on ESI-LTQ Orbitrap Hybrid mass

spectrometer. The molecular weights of all potential mono- and

oligosaccharides species that could be generated from incomplete

digestion of a PNAG sample with all possible acetylation patterns

are calculated. The corresponding m/z value for all these

molecules and their dehydration products are given here. This

table provides a complete list of all potential digestion products,

however only a subset of them might exist in the sample and would

show up on the spectrum.


Table S2 Validation of microarray data for identification of

mutants defective in responding to sPNAG. In-frame deletions of

the top candidate ORFs (and a few others) were generated and

their behavior was studied in the presence of 0.1 U/ml sPNAG.

Only deletion of rfaP, rfaQ, and rfaY impaired the biofilm

formation ability of the cells. Other abundant transposon insertion

events in different locations of the rfaQ-K operon presumably

caused their phenotype through polar effects or other indirect

interferences with proper biological functioning of rfaP, rfaQ, and

rfaY gene products. rfaF and rfaG deletion mutants showed

spontaneous surface-attachment activity in a sPNAG-independent

manner, as well. However, this phenotype was much weaker than

the sPNAG-induced behavior and was easily distinguishable from

it.


Table S3 Effect of Ca2+ concentration on cellular response to

sPNAG. Divalent cations are postulated to be involved in

maintaining outer membrane stability by cross-linking adjacent

LPS molecules through their phosphate groups ([10] in Text S1).

Consequently, increasing concentration of divalent cations should

mask the accessible phosphate groups exposed on the LPS

structure. Therefore, we reasoned that increasing concentrations

of divalent cations should interfere with any presumable

electrostatic interaction between phosphate groups and sPNAG.

To test this, wild-type cells were exposed to sPNAG in LB

supplemented with different concentrations of Ca2+. As shown

here, response to sPNAG is lost in Ca2+ concentrations higher

than 100 mM.

Found at: doi:10.1371/journal.ppat.1000432.s015 (0.03 MB

DOC)

Table S4 Validation of microarray data for identification of

suppressor mutations reverting the phenotype of DrfaY cells. In-

frame deletions of the top candidate ORFs (and a few others)

were generated in the DrfaY background and their behavior was

studied in the presence of 0.1 U/ml sPNAG. As shown here,

deletion of many of the LPS biosynthetic genes suppressed the

rfaY deletion phenotype. Based on the microarray results,

mutants with transposon insertions within or in the vicinity of

znuA and znuC, subunits of zinc uptake transporter of the inner

membrane ([11] in Text S1), were significantly enriched in our

selection. However, we did not observe any phenotype upon

deletion of any of those genes in the DrfaY background. We also

deleted zur, the transcriptional repressor of the znuBC and znuA

transcription units ([11] in Text S1) to explore the possibility of

the transposon disrupting the repressor binding site and

enhancing the expression of these genes, but no phenotype was

observed in that case either. Three of the genes involved in E. coli

type 1 fimbrium (pilus) biosynthesis, fimA, fimB, and fimE, also

acquired high scores in our selection. However, since type 1

fimbrium can enhance nonspecific adhesion to biotic and abiotic

surfaces ([12,13] in Text S1), we did not focus on those mutants.

In case of rfbD and rfbC mutants, the suppression effect was

stronger on glass surface, compared to plastic.


Table S5 Biological function of the rfaY suppressors associated

with amino acid decarboxylase systems (acid tolerance system)

identified in transposon insertion or over-expression libraries. E.

coli has evolved multiple acid resistance systems. Two of these

systems are based upon decarboxylation of glutamate or arginine

amino acids at the expense of a proton, which increases

intracellular pH ([14] in Text S1). Deletion of adiY, the positive

regulator of the arginine decarboxylase system, can suppress the

rfaY deletion phenotype. On the other hand, over-expression of

argR, the repressor of arginine biosynthesis, reverts the DrfaY

phenotype as well, and arginine is required for maintaining

activity of arginine decarboxylase system. These data suggest that

inactivation of the arginine decarboxylase system positively

contributes to sPNAG-based biofilm formation in the DrfaY

background. The same reasoning can be extended to gadWYX

over-expression. gadW and gadX are dual regulators of glutamate

decarboxylase system. Depletion of the genes belonging to ‘‘Acid

Stress Response Up-regulated’’ category in the microarray results

of the over-expression library (Figure S5) suggests that gadWYX

over-expression exerts its suppressor effect through down-regula-

tion of acid resistance, or more specifically glutamate decarbox-

ylase system. This is in agreement with the predicted behavior of

arginine decarboxylase system as well. There is also previous

evidence that gadE, the transcriptional activator of the acid

resistance system in E. coli, can up-regulate transcription of the

rfaQ-K operon ([15] in Text S1). We have also seen that

transcription of the rfaQ-K operon is decreased upon over-

expression of gadXYW (Figure 6C), suggesting that gadXYW over-

expression indirectly contributes to biofilm formation in the DrfaY

background through down-regulation of the acid tolerance system.

This presumably happens through structural modifications

imposed on LPS.




DOC)

Table S6 Surface antigen profile of clinical isolates of E. coli used

in this study. Eleven clinical isolates of E. coli were analyzed for

presence or absence of O-antigen and K1 capsule along with their

ability to respond to sPNAG. Among these 11 clinical strains were

7 urinary tract infection (UTI) isolates, including UTI-E, -G, -H, -

J, -P, -R, and -U, and 4 blood isolates from patients in neonatal

intensive care units (NICU), including NICU-2, -4, -10, and -12.

Presence or absence of O-antigen was determined after running

the LPS samples on SDS-PAGE gel and silver staining (Figure S6).

K1 capsule production was reported either by sensitivity to

bacteriophage E ([16] in Text S1) or based on previous data ([4] in

Text S1). NICU-2 was E-phage resistant, suggesting that it is

either producing a different variant of K1 capsule or has

completely lost its ability to produce K1 capsule due to some

spontaneous mutation. Endogenous PNAG production was

reported based on previous data ([4] in Text S1). All these strains,

except for UTI-H, were capable of expressing the pga locus, with

UTI-E having relatively weaker expression. Among these strains, 7

formed biofilms in the presence of sPNAG. Two of the UTI

isolates (UTI-U and UTI-P), both capable of responding to

sPNAG, were O-antigen2. Among the O-antigen+ strains, there

was no significant correlation between the abundance or length of

high molecular weight versions of O-antigen and the ability to

respond to sPNAG. Among NICU isolates, three K1+ NICU

isolates were also capable of responding to sPNAG.


DOC)

Table S7 Bacterial strains used in this study. *: csrA has been

reported to be conditionally essential in MG1655 ([17] in Text S1)

and csrA deletion mutants of MG1655 were reported to be capable

of growth on LB only after inactivation of the glgCAP operon.

However, we managed to delete the entire csrA ORF in our strain.

This discrepancy could be explained either by the differences that

might exist between the genetic backgrounds of the strains (which

could potentially make csrA deletion permissive in one) or

acquisition of suppressor mutations that allowed csrA deletion.

However, we found the growth rate of the mutant to be

significantly slower than the wild-type strain.


Table S8 Plasmids and bacteriophages used in this study.


Text S1 References in supporting information.


DOC)

Video S1 Time-lapse microscopy of wild-type MG1655 cells in

presence of 0.1 U/ml of sPNAG in LB. Images were acquired

every 30 seconds for the first three hours of exposure to sPNAG.


WMV)

Video S2 Time-lapse microscopy of wild-type MG1655 cells in

LB, in the absence of sPNAG. Images were acquired every

2 minutes for the first 6 hours of the experiment.


WMV)

Video S3 Time-lapse microscopy of UV-irradiated wild-type

MG1655 cells in presence of 0.1 U/ml of sPNAG in LB. Images

were acquired every 2 minutes for the first 6 hours of exposure to

sPNAG. The viable count of the cells after UV irradiation was

found to be zero.

Found at: doi:10.1371/journal.ppat.1000432.s024 (8.8 MB AVI)

Acknowledgments

We thank all members of the Tavazoie laboratory for helpful discussions

and critical reading of the paper. We are particularly grateful to Hany

Girgis for his help with transposon mutagenesis techniques and library. We

thank Margaret Bisher and Joseph Goodhouse for assisting with the

electron microscopy. We would also like to thank Ileana Cristea and Saw

Kyin for their help with mass spectrometry. We are grateful to Joseph Sklar

for the kind gift of the over-expression plasmid pool. We are indebted to

Gerald Pier, Eric Vimr, Miguel Valvano, and Gisela Storz for sending us S.

aureus PNAG sample, plasmids, and strains. Dispersin B was a kind gift

from Kane Biotech Inc.

Author Contributions

Conceived and designed the experiments: SA ST. Performed the

experiments: SA HG. Analyzed the data: SA HG. Wrote the paper: SA

HG ST.

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